Glutathione-dependent redox status of frataxin-deficient cells in a yeast model of Friedreich's ataxia.
ABSTRACT Friedreich's ataxia is a neurodegenerative disease caused by reduced expression of the mitochondrial protein frataxin. The main phenotypic features of frataxin-deficient human and yeast cells include iron accumulation in mitochondria, iron-sulphur cluster defects and high sensitivity to oxidative stress. Glutathione is a major protective agent against oxidative damage and glutathione-related systems participate in maintaining the cellular thiol/disulfide status and the reduced environment of the cell. Here, we present the first detailed biochemical study of the glutathione-dependent redox status of wild-type and frataxin-deficient cells in a yeast model of the disease. There were five times less total glutathione (GSH+GSSG) in frataxin-deficient cells, imbalanced GSH/GSSG pools and higher glutathione peroxidase activity. The pentose phosphate pathway was stimulated in frataxin-deficient cells, glucose-6-phosphate dehydrogenase activity was three times higher than in wild-type cells and this was coupled to a defect in the NADPH/NADP(+) pool. Moreover, analysis of gene expression confirms the adaptative response of mutant cells to stress conditions and we bring evidence for a strong relation between the glutathione-dependent redox status of the cells and iron homeostasis. Dynamic studies show that intracellular glutathione levels reflect an adaptation of cells to iron stress conditions, and allow to distinguish constitutive stress observed in frataxin-deficient cells from the acute response of wild-type cells. In conclusion, our findings provide evidence for an impairment of glutathione homeostasis in a yeast model of Friedreich's ataxia and identify glutathione as a valuable indicator of the redox status of frataxin-deficient cells.
- SourceAvailable from: Lidia Cova[show abstract] [hide abstract]
ABSTRACT: Friedreich ataxia is a progressive neurodegenerative disorder caused by loss of function mutations in the frataxin gene. In order to unravel frataxin function we developed monoclonal antibodies raised against different regions of the protein. These antibodies detect a processed 18 kDa protein in various human and mouse tissues and cell lines that is severely reduced in Friedreich ataxia patients. By immunocytofluorescence and immunocytoelectron microscopy we show that frataxin is located in mitochondria, associated with the mitochondrial membranes and crests. Analysis of cellular localization of various truncated forms of frataxin expressed in cultured cells and evidence of removal of an N-terminal epitope during protein maturation demonstrated that the mitochondrial targetting sequence is encoded by the first 20 amino acids. Given the shared clinical features between Friedreich ataxia, vitamin E deficiency and some mitochondriopathies, our data suggest that a reduction in frataxin results in oxidative damage.Human Molecular Genetics 11/1997; 6(11):1771-80. · 7.69 Impact Factor
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ABSTRACT: Friedreich's ataxia (FRDA) is an autosomal recessive, degenerative disease that involves the central and peripheral nervous systems and the heart. A gene, X25, was identified in the critical region for the FRDA locus on chromosome 9q13. The gene encodes a 210-amino acid protein, frataxin, that has homologs in distant species such as Caenorhabditis elegans and yeast. A few FRDA patients were found to have point mutations in X25, but the majority were homozygous for an unstable GAA trinucleotide expansion in the first X25 intron.Science 04/1996; · 31.03 Impact Factor
Chapter: Iron and Friedreich ataxia[show abstract] [hide abstract]
ABSTRACT: Friedreich ataxia is due to insufficient levels of frataxin, a mitochondrial iron chaperone that shields this metal from reactive oxygen species (ROS) and renders it bioavailable as Fe II. Frataxin participates in the synthesis of iron-sulfur clusters (ISCs), cofactors of several enzymes, including mitochondrial and cytosolic aconitase, complexes I, II and III of the respiratory chain, and ferrochelatase. It also plays a role in the maintenance of ISCs, in particular for mitochondrial aconitase. A role of frataxin in heme synthesis has been postulated, but is controversial. Insufficient frataxin leads to deficit of ISC enzymes and energy deficit. Iron levels increase in mitochondria. Oxidative stress may result from respiratory chain dysfunction and from direct reaction between iron and ROS. Stress pathways are activated that may lead to apoptosis or other forms of cell death. The basis for the selective vulnerability of specific neurons, like sensory neurons, is still unknown.12/2005: pages 143-146;
Glutathione-dependent redox status of
frataxin-deficient cells in a yeast model
of Friedreich’s ataxia
Franc ¸oise Auche `re?, Renata Santos, Sara Planamente, Emmanuel Lesuisse and
Laboratoire d’Inge ´nierie des Prote ´ines et Contro ˆle Me ´tabolique, De ´partement de Biologie des Ge ´nomes,
Institut Jacques Monod, UMR 7592, CNRS, Universite ´s Paris 6 and 7, 2 Place Jussieu, Tour 43,
75251 Paris Cedex 05, France
Received February 19, 2008; Revised May 16, 2008; Accepted June 16, 2008
Friedreich0s ataxia is a neurodegenerative disease caused by reduced expression of the mitochondrial
protein frataxin. The main phenotypic features of frataxin-deficient human and yeast cells include iron
accumulation in mitochondria, iron-sulphur cluster defects and high sensitivity to oxidative stress.
Glutathione is a major protective agent against oxidative damage and glutathione-related systems participate
in maintaining the cellular thiol/disulfide status and the reduced environment of the cell. Here, we present the
first detailed biochemical study of the glutathione-dependent redox status of wild-type and frataxin-deficient
cells in a yeast model of the disease. There were five times less total glutathione (GSH1GSSG) in frataxin-
deficient cells, imbalanced GSH/GSSG pools and higher glutathione peroxidase activity. The pentose phos-
phate pathway was stimulated in frataxin-deficient cells, glucose-6-phosphate dehydrogenase activity was
three times higher than in wild-type cells and this was coupled to a defect in the NADPH/NADP1pool.
Moreover, analysis of gene expression confirms the adaptative response of mutant cells to stress conditions
and we bring evidence for a strong relation between the glutathione-dependent redox status of the cells and
iron homeostasis. Dynamic studies show that intracellular glutathione levels reflect an adaptation of cells to
iron stress conditions, and allow to distinguish constitutive stress observed in frataxin-deficient cells from
the acute response of wild-type cells. In conclusion, our findings provide evidence for an impairment of
glutathione homeostasis in a yeast model of Friedreich’s ataxia and identify glutathione as a valuable
indicator of the redox status of frataxin-deficient cells.
Friedreich0s ataxia (FA) is a neurodegenerative disease charac-
terized by progressive neurodegeneration and cardiomyopathy.
It is an inherited autosomal recessive disorder caused by
reduced expression of the mitochondrial protein frataxin (1,2).
The major phenotypic features of frataxin-deficient human
and yeast cells include iron accumulation in mitochondria,
iron–sulphur cluster defects and high sensitivity to oxidative
stress (reviewed in 3). This raises the question of the status of
the cellular defences against oxidative injury in FA cells.
There are preliminary experiments suggesting that patients
with FA suffer a disturbance of glutathione homeostasis and
modifications of glutathione-dependent antioxidant defences
Thiol (-SH)-containing antioxidants, including cystein and
glutathione, make a large contribution to the protection
against oxidative damage; they do so by reacting with reactive
oxygen species (ROS). In yeast, the thiol-dependent response
to oxidative stress consists of the parallel glutathione/
glutaredoxin and thioredoxin pathways which are both
required under aerobic and anaerobic conditions (reviewed
(GSH/GSSG) is the most abundant low-molecular-weight non-
protein thiol in eukaryotic cells (11,12). It is involved in a
variety of cellular functions: protection against oxidative
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Human Molecular Genetics, 2008, Vol. 17, No. 18
Advance Access published on June 18, 2008
by guest on June 5, 2013
damage, the maintenance of the mitochondrial structure and
membrane integrity, cell differentiation and development
(13,14). Due to its low redox potential (E00¼2240 mV) and
its high cellular concentration (1–10 mM), glutathione is a
major redox buffer in thiol-based redox systems (7). It is also
a strong nucleophile and conjugates with ROS and with a
range of electrophilic compounds and numerous xenobiotics
(15). Its unusual g-glutamyl peptide bond results in it having
substantial resistance to proteolytic degradation. GSH depletion
results in a particular defect in the maturation of iron–sulphur
proteins, leading to iron accumulation in the cells (16,17).
The reduced sulphydryl group in glutathione (GSH), when
oxidized, produces a disulphide, or ‘oxidized glutathione’
(GSSG), and functions as a source of reducing equivalents
to reduce cellular disulphide bonds, often in conjunction
with glutaredoxins (8,18). Oxidants interfere with a range of
cellular components and cause accumulation of ROS and/or
alter the GSH/GSSG ratio, and as a consequence changes in
GSH concentration directly reflect intracellular redox changes.
The levels of intracellular-reduced glutathione are maintained
by strict regulation of a combination of glutathione-dependent
enzymatic systems that balance its rate of synthesis and
reduction. Glutathione reductases restore and maintain physio-
logical GSH/GSSG balances under both stressed and unstressed
conditions by reducing GSSG in a NADPH-dependent reaction.
The response to oxidative stress also involves GSH biosynthesis
enzymes, and peroxide-eliminating glutathione peroxidases and
glutaredoxins (19). In addition, glutathione-S-transferases play
a protective role by attaching GSH to biomolecules, targeting
them for export from the cell (20).
Yeast also contains a thioredoxin system, including thiore-
doxin and thioredoxin reductase, which functions as a protec-
tion against ROS generated during respiratory metabolism
(21–23). Thioredoxins, like glutaredoxins, are small heat-
stable oxidoreductases containing two conserved cystein resi-
dues in their active site. The oxidized disulphide form of
thioredoxin is reduced directly by NADPH and thioredoxin
reductase, whereas glutaredoxin is reduced by glutathione
with NADPH as the electron donor (8). The thioredoxin
system involves both cytosolic and mitochondrial thioredoxin
peroxidases (24,25). However, in Saccharomyces cerevisiae,
the thioredoxin and GSH/glutaredoxin systems are not func-
tionally redundant. Glutathione is involved in critical func-
tions of redox regulation not shared with the thioredoxin
pathway, and works on Dgsh1 mutants suggest that GSH is
important for the maintenance of the mitochondrial genome
(26,27). S. cerevisiae
(lacking GSH1) or altered in their GSH redox state are
unable to grow in the absence of GSH and are less tolerant
than wild-type to a wide range of stress conditions induced
by peroxides and the superoxide anion, and to the toxic
products of lipid peroxidation; they also undergo apoptosis
more than their parental cells (28–31). Depletion of mitochon-
drial and cytosolic glutathione is associated with increased
generation of ROS, defects in iron–sulphur cluster assembly
and rapid loss of mitochondrial function (16,17,19,26,32,33).
Inversely, administration of exogenous glutathione is capable
of rescuing cells from accelerated aging (34).
Abnormal glutathione homeostasis has been described in
various neurodegenerative diseases, and during cell aging and
including increased degradation, conjugation, oxidation, efflux
and biosynthesis decrease(12,13,36,37).Yeast is a good eukary-
and biochemical tools are available for manipulations. Partly
because the significance of glutathione in neurodegeneration is
becoming increasing evident and because of potential appli-
cations as a clinical marker of oxidative stress (38), we studied
glutathione-dependent redox status in a S. cerevisiae model of
Friedreich’s ataxia, and of the wild-type.
We show that GSH/GSSG levels in frataxin-deficient cells
are low, and this is associated with high activities of thiol-
dependent enzymes including glutathione peroxidases. The
HPLC analysis of NADPH/NADPþpools demonstrated a
relation between the glutathione-redox status and a strong
stimulation of the pentose phosphate pathway. The changes
in thiol-dependent antioxidants were associated with modifi-
cations of gene transcription.
We also report the glutathione-dependent response to iron
excess conditions. Dynamic studies show that intracellular
glutathione levels in wild-type and mutant cells reflect
adaptation to iron excess conditions. Addition of excess iron
to cultures of the mutant cells resulted in GSH levels similar
to those in wild-type cells. Our findings reveal that the hyper-
sensitivity of frataxin-deficient cells to oxidative stress is
associated with modifications of the glutathione-dependent
antioxidant defences and possibly of glucose metabolism.
They are also consistent with there being substantial physio-
logical interaction between iron and glutathione pathways.
Total glutathione levels (GSH1GSSG) are decreased
in frataxin-deficient cells
Total (GSHþGSSG) and oxidized (GSSG) glutathione levels
were measured (Fig. 1). Wild-type cells (strain S150-2B)
grown in minimum YNB medium under normal aerobic con-
ditions have a redox ratio (GSH/GSSG) of ?6/1 (Fig. 1) indicat-
ing that most of the intracellular free glutathione is in a reduced
(GSH) form; this prevents the toxic formation of mixed
disulphur-protein products, as previously described (10,39).
Frataxin-deficient cells contained less total glutathione, with
87.2+19.2 nmoles glutathione/mg protein, rather than the
173.4+23.5 nmoles glutathione/mg protein in wild-type cells
(Fig. 1). In addition, the redox balance (GSH/GSSG) was
shifted towards the oxidized form GSSG, reflecting a depletion
of the reduced form of glutathione such that there were similar
amounts of the reduced and oxidized form in the mutant cells
(theGSH/GSSGratio was 0.8). The GSH/GSSGratio, therefore,
sical exposure to stress conditions, in which oxidized GSSG
levels are elevated in parallel with a decrease in intracellular
GSH levels (10,29). Similar results were obtained when cells
were grown on a rich YPD-medium and with strain YPH499,
showing that the overall glutathione defect was not strain
dependent (data not shown).
dria isolated from frataxin-deficient cells (11.6+1.2 nmoles
Human Molecular Genetics, 2008, Vol. 17, No. 18 2791
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glutathione/mg protein in wild-type cells and 40.3+5.4 nmoles
glutathione/mg protein in Dyfh1 cells). Interestingly, the distri-
bution of glutathione pools between cytosol and mitochondria
was abnormal in the mutant. Most glutathione (93.3%) in wild-
type cells is on the cytosol, but, in frataxin-deficient cells,
46.2% of total glutathione was found inside the isolated mito-
chondria. S. cerevisiae mitochondria cannot synthesize gluta-
thione, so these findings suggest import of glutathione into the
mitochondrial matrix from the cytosol in response to the oxi-
dative stress associated with the frataxin deficiency.
Glutathione peroxidase activity is enhanced in
Glutathione peroxidases (Gpx) are the main peroxide detoxify-
ing enzymes in S. cerevisiae, and act by catalysing the break-
down of H2O2and larger hydroperoxides using GSH as a
reductant (40). These enzymes are required for cellular protec-
tion against lipid peroxidation. Three gene products (Gpx1,
Gpx2 and Gpx3) with significant homology (30, 41 and
40%, respectively) to mammalian peroxidases have been
identified from the yeast genome (41–45). We assayed gluta-
thione peroxidase activities in crude cell extracts and isolated
mitochondria (Fig. 2). Gpx activity was much higher in
frataxin-deficient cells than wild-type cells (specific activities
of 995.9+75.8 units/mg and 2424.3+107.1 units/mg in
wild-type and frataxin-deficient cells), which confirm the
severe oxidative stress status of the mutant cells. The specific
activity in isolated mitochondria was similarly higher in the
mutant than wild-type cells (Fig. 2). The lower specific
activity in isolated mitochondria than in cell extracts was pre-
sumably due to most of the Gpx being in the cytoplasm. Thus,
the Gpx activity in cytosol is 2.3 times higher in Dyfh1 than
wild-type cells and three times higher in mitochondria, con-
sistent with the difference in total glutathione levels and
with the thiol-dependent antioxidant defences being ‘switch-
on’ in frataxin-deficient cells.
The total reduced thiol groups are depleted in
S. cerevisiae frataxin-deficient cells
We assayed total reduced thiol groups in wild-type and mutant
cells (Fig. 3). Total thiol groups include free glutathione, free
thiol groups such as cystein, and protein-bound thiol groups.
Crude extracts of wild-type
24.7 nmoles of total thiols/mg of protein and those of frataxin-
deficient cells 83.0+11.6 nmoles total thiols/mg protein. This
difference is consistent with the severe stress status of the
frataxin-deficient cells and their lower total glutathione
content. Moreover, the respective amounts of reduced gluta-
thione (calculated from Fig. 1) and total reduced thiol
groups are in good agreement with glutathione being the
main non-protein thiol molecule in the cell (85.4%).
However, the amounts of reduced thiol groups in mitochondria
were similar in mutant and wild-type cells (27.1+2.7 nmoles
total thiols/mg protein, Fig. 3). These results could suggest a
difference in the distribution of thiol groups between cytosol
and mitochondria, or transport of these groups to the surface
of the cell. Indeed, when cells or tissues are subjected to oxi-
dative stress, export of GSSG has been observed and this
efflux appears to be a part of the protection of cells and
tissues against oxidative stress. To further explore this issue,
extracellular glutathione was assayed; there was no significant
difference between wild-type and frataxin-deficient cells, and
Figure 1. Total glutathione (GSH þ GSSG) and oxidized glutathione disul-
phide (GSSG) in wild-type and frataxin-deficient cells (strain S150-2B).
Total (black bars) and oxidized (grey bars) glutathione levels were determined
in crude cell extracts using the enzymatic recycling assay (see Materials and
Methods section), and normalized to the protein content determined by the
BCA protein assay. All data points in the figure represent means of at least
Figure 2. Glutathione peroxidase activity in wild-type and frataxin-deficient
cells (strain S150-2B). Specific glutathione peroxidase activities were
assayed using ter-butylhydroperoxide (t-BHP) as a substrate, in cell extracts
(left panel) and isolated mitochondria (right panel); values for wild-type
cells are in black and those for mutant cells in grey. All data points show
means of at least three determinations.
Figure 3. Total reduced thiol groups in wild-type and frataxin-deficient cells.
Total reduced thiol groups in cell extracts or isolated mitochondria were
5.50-dithiobis-2-nitrobenzoı ¨c acid (DTNB) into 5-thio-2-nitrobenzoı ¨c acid
(TNB) at 412 nm; values for wild-type cells are in black and those for
mutant cells in grey. All data points in the figure and the values listed are
expressed in nanomoles of total reduced thiols/mg of protein and are means
of at least three determinations.
of the conversionof
2792Human Molecular Genetics, 2008, Vol. 17, No. 18
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therefore no evidence of enhanced export of glutathione from
The pentose phosphate pathway is strongly stimulated
in S. cerevisiae frataxin-deficient cells
Most of the electron sources for the two glutathione/glutare-
doxin and thioredoxin systems are provided by NADPH
which, due to its low redox potential (E0¼ 2315 mV), can
act as the primary hydrogen donor for both systems. Indeed,
glutathione reductases maintain physiological GSH/GSSG bal-
ances by reducing GSSG in a NADPH-dependent reaction,
and the oxidized disulphide form of thioredoxin is reduced
directly by NADPH and thioredoxin reductase. We determined
relative NADPH and NADPþconcentrations in wild-type and
mutant cells by reverse-phase HPLC experiments (Table 1).
Under our experimental conditions, wild-type cells contained
120.9+8.3 nmoles NADPH/mg protein, and mutant cells
29.3+12.7 nmoles NADPH/mg protein. These results are in
good agreement with the higher glutathione peroxidase
activity observed in mutant cells (Fig. 2): GSSG is reduced
back to GSH by glutathione reductase using NADPH for
The NADPH/NADPþratios were 1.9 in wild-type cells and
0.6 in mutant cells, indicating a substantially lower pool of the
(Table 1). Depletion of reduced NADPH is characteristic of
severe oxidative stress conditions, which, for whatever
reason, cannot be counterbalanced by antioxidant systems in
the cells. Glucose-6-phosphate dehydrogenase and, to a
(IDP) are believed to be the major sources of NADPH used
by thiol-dependent peroxidases in the cytosol of S. cerevisiae
cells (46), so these findings suggest that the pentose phosphate
pathway is strongly perturbed in frataxin-deficient cells.
Glucose-6-phosphate dehydrogenase is the key enzyme of
the pentose phosphate pathway that is responsible for the
generation of NADPH and plays a role in the protection
against oxidative damage in eukaryotic cells (47,48). Specific
G6PDH activity was significantly higher in frataxin-deficient
in frataxin-deficient cells
(53.4+5.9 nmoles NADPH/min/mg protein) cells. This indi-
cates, as suggested by the NADPH/NADPþratios, that the
pentose phosphate pathway is stimulated in frataxin-deficient
cells. Indeed, when NADPH is being rapidly converted to
NADPþ, as in frataxin-deficient cells, the level of NADPþrises,
allosterically stimulating G6PDH to accelerate NADPH biosyn-
thesis. These observations are also in good agreement with pre-
vious work showing that G6PDH expression is enhanced by
oxidative stress induced by agents that specifically deplete the
intracellular glutathione pool (49,50).
Regulation of genes
detoxification in frataxin-deficient cells. We used real-time
quantitative PCR for transcriptional analysis of the genes
implicated in the glutathione-dependent systems of ROS
detoxification in wild-type and frataxin-deficient cells. The
results (Table 2) are in good agreement with the biochemical
data. The ratios of expression of the genes GSH1 and GSH2 in
frataxin-deficient (1.15) and wild-type cells (1.32) were
similar, consistent with the lower glutathione levels in
mutant cells being a consequence of pool redistribution and
not reduced biosynthesis. Transcription of the gene encoding
the high-affinity glutathione transporter OPT1 was twice as
high in frataxin-deficient cells, consistent with the decrease
in total glutathione (Table 2). However, transcription of
OPT1 has been reported to be repressed by exogenous GSH
but not regulated in a Dgsh1 mutant deficient in glutathione
synthesis (51). Our results present no evidence of a change
in transcription of the glutaredoxin gene family GRX1-5, nor
in that of the glutathione transferase GTT1-3 (Table 2).
Frataxin deficiency causes an oxidative stress and depletion
of glutathione and NADPH pools and induction of Gpx and
G6PDH activities. Accordingly, transcription of GPX2 and
ZWF1 genes was twice as high as wild-type in Dyfh1 mutant
cells (Table 2). However, the gene GLR1 encoding the gluta-
thione reductase that uses NADPH to reduce GSSG was unaf-
fected in frataxin-deficient cells (Table 2). These various
results show that the changes in enzymatic and thiol levels
are due to transcriptional and post-transcriptional modifi-
cations, and reveal the particular metabolic patterns in
involvedin thiol-dependent ROS
Glutathione redox status of frataxin-deficient S. cerevisiae
cells is modified in the presence of excess iron
In S. cerevisiae, deletion of the frataxin homolog YFH1 results
in a 10-fold increase in iron in mitochondria and increased
ROS production (52,53). It has been suggested that this iron
excess is responsible both for ROS production by the Fenton
reaction and for secondary phenotypes such as aconitase
deficiency and mitochondrial DNA loss. It has also been
suggested that the glutathione-dependent redox status of the
cells and iron homeostasis are linked. GSH depletion causes
a specific defect in the maturation of iron–sulphur proteins,
leading to iron accumulation in the mitochondria (17). In
addition, deficiency in the mitochondrial glutaredoxin Grx5
associates defective activity of mitochondrial iron–sulphur
proteins and cellular iron accumulation (16).
We grew cells in presence of various concentrations of
radioactive55Fe and followed iron accumulation (Fig. 4). As
Table 1. NADPH/NADPþlevels and glucose-6-phosphate dehydrogenase
(G6PDH) activities in wild-type and frataxin-deficient cells
NADPH (nmoles/mg protein)
Glucose 6-phosphate dehydrogenase specific
activity (nmoles NADPH/min/mg protein)
After acidic or alkaline extraction of the cellular extracts, NADPH and
NADPþlevels were assayed by reverse phase HPLC. Products were
monitored spectrophotometrically at 260 nm and quantified by integration
of the peak absorbance area, employing a calibration curve of increasing
concentrations of NADPH and NADPþ. Glucose-6-phosphate
dehydrogenase activities were assayed by following the increase of
absorbance at 340 nm due to the conversion of NADPþto NADPH.
Wild-type cells and frataxin-deficient cells were grown in YNB-glucose
minimum medium and all values are means+SD of at least three
Human Molecular Genetics, 2008, Vol. 17, No. 182793
by guest on June 5, 2013
expected, iron accumulation was much higher in Dyfh1 than
wild-type cells, and was proportional to the amount of iron
added to the culture medium. At 100 mM iron, the highest con-
centration tested, corresponding to a 100-fold more than in
standard YNB medium, mutant cells accumulated 614.5+
104.8 pmoles Fe/OD600nm, and wild-type cells accumulated
105.9+27.7 pmoles Fe/OD600nm(Fig. 4). Consequently, we
studied the glutathione-dependent redox status of wild-type
and mutant cells grown in the presence of a 100 mM iron.
Thetotal glutathione (GSHþGSSG)
(Fig. 5A) in mutant cells cultivated in the presence of an
excess iron was similar to that in wild-type cells, and nearly
double than in mutant cells cultured on YNB medium. The
presence of excess iron was associated with a decrease of
the oxidized disulphide GSSG; 48.4+2.3 nmoles GSSG/mg
protein and 18+5.7 nmoles GSSG/mg protein for Dyfh1 cul-
tivated in minimum and iron–supplemented YNB medium,
respectively (data not shown). In the presence of excess
iron, there was also a 10-fold increase of the GSH/GSSG
ratio, suggesting a return to non-stress redox conditions. At
the transcriptional level, the raise in total glutathione cannot
be easily attributed to an increased synthesis, because at
least GSH1 and GSH2 transcription was not affected by iron
addition (Table 2). Interestingly, the expression of the high-
affinity glutathione transporter encoding gene OPT1 was not
modified and remained up regulated which suggested that
cells still sense the need of external glutathione in the presence
of iron. As shown in Figure 5B, glutathione peroxidase activi-
ties were decreased in the presence of an excess iron and were
similar to the ones observed for wild-type cells. In agreement
with reduced Gpx activity, GPX2 transcription was lowered in
frataxin-deficient cells cultured with excess iron but the gene
remained induced compared to wild-type cells (Table 2).
The consequences of the presence of the iron chelator BPS
(bathophenanthrolin disulphonic acid) were tested, but the
glutathione-dependent redox status and Gpx activity were
unaffected (Fig. 5A and B).
As shown in Figure 5C, G6PDH activity in mutant cells cul-
tivated with an excess iron was 102.8+3.9 nmoles NADPH/
min/mg protein, lower than the 155.5+20.1 for mutant cells
grown in standard minimum medium. However, in presence of
an excess iron, G6PDH activity of frataxin-deficient cells was
Table 2. Analysis of the expression of genes implicated in the glutathione-dependent response to oxidative stress
GenePrimers (50g30) Relative expression Dyfh1/wt
YNBYNB þ Fe
Cells were cultivated on YNB-medium, supplemented or not with 100 mM iron citrate, to exponential phase. Relative expression was calculated using
the equation described by (79). Sequences of primers used are depicted, the first is the forward and the second the reverse (nd, not determined).
2794 Human Molecular Genetics, 2008, Vol. 17, No. 18
by guest on June 5, 2013
still higher than the one observed in wild-type cells (Fig. 5C).
As shown in Table 2, ZWF1 gene remains activated in the pre-
sence of iron, which confirms that the adaptative response to
iron stress conditions can be attributed to metabolic choices
of the cells. The major change in transcription in the presence
of iron was the induction of mitochondrial glutathione-S-trans-
ferase GTT2, the main transferase involved in cadmium and
oxidative stress resistance and adaptation (54,55). These
results show that the glutathione-dependent redox status of
mutant cells is strongly dependent on the abundance of iron
in the environment and could reflect an adaptative response
of the cells to iron-dependent stress conditions.
Iron-dependent glutathione redox status reflects an
adaptative response to excess iron conditions
In response to stress conditions, yeast cells up regulate the
synthesis of a number of protective molecules including com-
ponents of both the GSH/glutaredoxin and the thioredoxin
systems (19,29). This forms the basis of an inducible adapta-
tive response, in which, for example, cells treated with a
low dose of oxidant can adapt to become resistant to a sub-
sequent stress and otherwise lethal treatment. In the following
experiments, wild-type and mutant cells were cultured in
minimum YNB medium until reaching the exponential phase
of growth: aliquots were treated with excess iron, and other
were transferred to an iron-depleted culture medium to
create iron-deficient conditions. Thiol-dependent antioxidants,
including glutathione concentrations and Gpx activities, were
followed for up to 2 h.
There was no change in total glutathione pools or Gpx
activity for wild-type or mutant cells when shifted, after fil-
tration, to the same YNB medium (Figs. 6A and 7A). This
control was very important to check that the filtration
process did not constitute a stress for the cells, and that any
changes in glutathione levels or enzymatic activities detected
could be attributed to the changes in the culture conditions.
For wild-type cells, the presence of an excess iron in
the culture medium resulted in a rapid decrease in the
total glutathione concentration (Fig. 6B), from 173.4+
23.5 nmoles glutathione/mg protein at baseline to 84.3+
8.7 nmoles glutathione/mg protein after 1 min of treatment.
Moreover, the calculated GSH/GSSG ratio was 6.0 at baseline
and 6.3 after 1 min with no subsequent change. In parallel, as
increased after 1 min in the presence of an excess iron
(Fig. 7B), followed by a return to normal conditions after
10 min. This suggests an acute response of wild-type cells to
iron stress conditions, followed by and adaptation and equili-
bration after 5 min and a return to non-stress conditions.
Repeat measurements, 120 min after the start of treatment
(data not shown), did not reveal any further changes in the
glutathione-dependent redox status of these cells, showing
adaptation of wild-type cells to the new culture conditions
was complete within 10 min.
Figure 4. Iron accumulation in wild-type and frataxin-deficient cells as a func-
tion of iron concentration. Iron accumulation in cells was measured after over-
night culture in minimum medium with 1–100 mM55Fe(II)-citrate. Values for
wild-type cells were in black and those for mutant cells in white. All data
points in the figure are means of at least three determinations.
Figure 5. Effect of iron on the glutathione-dependent redox status of wild-type
and frataxin-deficient cells. Cells were cultivated on YNB-medium, with and
without supplementation with 100 mM iron citrate or 30 mM BPS; values for
wild-type cells are in black, and mutant cells in grey and all are means of three
or more determinations. (A) Total glutathione levels; (B) glutathione peroxi-
dase (Gpx) specific activities. (C) Glucose-6-phosphate dehydrogenase
Human Molecular Genetics, 2008, Vol. 17, No. 182795
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In contrast, excess iron had little or no effect on glutathione
levels or on GSH/GSSG ratios in Dyfh1 cells (Fig. 6B).
However, Gpx specific activity decreased from 2424.3+
107.1 units/mg protein at baseline to 1645+138.6 units/mg
protein after 1 min, and returned to the initial value after
10 min (Fig. 7B). Thus, wild-type and mutant cells respond
very differently to excess iron and show the fundamental
difference between an acute and a constitutive response to
iron excess conditions.
We found that iron deficiency constituted a stress for wild-
type cells. Indeed, total glutathione levels increased after
1 min (Fig. 6C), then decreased to a value lower than baseline
value observed for wild-type cells. This observation suggests
that the cells have only a poor capacity to continue their adap-
tative response in conditions of severe iron deficiency.
However, there was no significant change in Gpx activity in
wild-type cells after exposure to an iron-depleted Bio101
culture medium (Fig. 7C). In contrast, Dyfh1 cells were not
sensitive to iron deficiency (Figs. 6C and 7C), which confirms
the opposite adaptative mechanisms of wild-type and constitu-
tively stressed mutant cells when subjected to iron excess
Figure 6. Glutathione-dependent adaptative response to iron stress conditions.
Cells were cultivated in YNB-medium to exponential phase, isolated by
filtration and shifted to different media: (A) YNB medium; (B) YNB
medium þ 100 mM iron citrate; (C) YNB-Fe (iron-depleted) medium.
Samples were taken 0, 1, 5, 10 and 15 min after cell filtration, and total
glutathione (GSHþGSSG) determined by the recycling enzymatic assay.
Values for wild-type cells are in black, and for frataxin-deficient cells
in grey, and all data points in the figure are means+SD of at least three
Figure 7. Glutathione-peroxidase adaptative response to iron stress conditions.
Cells were cultivated in YNB-medium to exponential phase, isolated by fil-
tration and shifted to: (A) YNB medium; (B) YNB medium þ 100 mM iron
citrate; (C) YNB-Fe (iron-depleted) medium. Samples were taken 0, 1, 5,
10 and 15 min after cell filtration, and assayed for specific glutathione peroxi-
dase activity, using ter-butylhydroperoxide (t-BHP) as a substrate. Values for
wild-type cells are in black, and those for frataxin-deficient cells in grey, and
all data are means+SD of at least three determinations.
2796Human Molecular Genetics, 2008, Vol. 17, No. 18
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We exploited the experimental advantages of S. cerevisiae to
conduct a detailed study of the glutathione-dependent cellular
defences of frataxin-deficient cells. The tripeptide glutathione
is an important molecule in the protection of yeast cells
against oxidative stress (9,10,19,28,30). In the first part of
this work, we compared the glutathione-dependent redox
status of wild-type and frataxin-deficient cells, and then we
extended the study to explore the adaptative response of
cells to iron-dependent stress conditions.
Frataxin-deficient cells are characterized by a deficiency of
total glutathione levels, associated with a shift of the GSH/
GSSG ratio towards the oxidized disulphide form GSSG
(Fig. 1). This is consistent with the role of GSH as both a free
radical scavenger and a cofactor for various antioxidant
enzymes, including glutathione peroxidases, glutathione-S-
transferases and glutaredoxins. The GSH/GSSG ratio reflects
the reducing conditions inside the cell, so this observation
agrees with classical exposure to stress conditions, where oxi-
dized GSSG levels are elevated and intracellular GSH levels
reduced. Total reduced thiol groups were also less abundant in
the mutant than wild-type (Fig. 3), confirming the abnormal
levels of GSH1 and GSH2, encoding the enzymes involved in
the glutathione biosynthesis pathway, are not modified in
frataxin-deficient cells (Table 2). Therefore, the low total gluta-
but to a consumption of glutathione associated with stress con-
ditions. Microarray data (not shown) do not reveal a change in
ECM38 (encoding g-glutamyl-transpeptidase). Also YCF1
(encoding vacuolar GSH transporter, Table 2) transcription is
unaffected so increased glutathione degradation in vacuoles
does not seem to be responsible for the low GSH concentration.
most likely not due to an abnormality in the activity of this
S. cerevisiae contains three glutathione peroxidases (Gpx1,
Gpx2 and Gpx3) which catalyse the breakdown of H2O2and
larger hydroperoxides using GSH as a reductant (30,40–43).
Two members of the glutaredoxins family, Grx1 and Grx2,
are also active as glutathione peroxidases (56). A significant
activation of the glutathione peroxidase pathway to ?150% of
control activity, associated with high GSSG levels, has been
described in lymphoblasts from FA patients (4). This is
similar to the two to three times higher Gpx activity in
mutants than wild-type S. cerevisiae cells (Fig. 2). Glutathione
peroxidases reduce H2O2and other organic hydroperoxides to
the corresponding alcohol, using the reducing power provided
by GSH. The higher Gpx activity is also consistent with obser-
vations that oxidized glutathione levels are significantly
higher in mutants, with a GSH/GSSG ratio indicative of a
stressed status of the cells. Gpx activity in mitochondria was
alsohighbutmostofthe enzymatic activitywas inthe cytosolic
fraction. Our transcription data indicate that GPX2 seems to be
the only gene controlled by Yap1 to be induced in mutant cells
tutive adaptation of Dyfh1 to oxidative stress conditions.
Glutathione is usually found in the cytosol, vacuole and
other compartments including the endoplasmic reticulum,
nucleus and mitochondria; it can be assimilated from the
medium by yeast cell-surface transporters (57). Under our
experimental conditions, glutathione was mainly found in
the cytosol in wild-type cells. There was 4-fold more total glu-
tathione in the mitochondria of mutant than wild-type cells
(Fig. 1), indicating a different distribution of the glutathione.
However, glutathione cannot be synthesized inside yeast mito-
chondria and must be imported to the mitochondrial matrix
from the cytosol. Our results suggest an import of glutathione
to the mitochondria under the stress conditions induced by fra-
taxin deficiency. This hypothesis is supported by the observed
induction of the OPT1, gene coding for glutathione transporter
in response to a decrease of intracellular total glutathione
levels (Table 2).
We also tested whether glutathione was exported from the
cells. GSH can be found outside the cells, but generally in
small amounts, and this extracellular glutathione is though to
function in detoxification processes and provide protection
against oxidative injury. Indeed, in response to an oxidative
challenge, oxidized GSSG, and protein-bound and extracellu-
lar glutathione are all elevated in parallel with the decrease of
intracellular GSH levels (29). However, under our conditions,
no glutathione excretion to the culture medium was detected
within 48 h. Furthermore, measurements of extracellular gluta-
thione in mutant cells provided no evidence of export of glu-
tathione out the cells.
FA patients have abnormally low free glutathione concen-
tration, high glutathione transferase (GST) activity, associated
with a significant increase of glutathione bound to haemo-
globin in erythrocytes (5,6), suggesting that protein glutathio-
nylation is a response to stress conditions induced by frataxin
deficiency. Indeed, cystein is the most easily oxidized residue
in proteins, resulting in intermolecular protein cross-linking
and enzyme inactivation (58). Such irreversible oxidation
events can be prevented by protein-S-thiolation in which
protein SH groups react with low molecular weight thiols
and particularly GSH (glutathionylation) (59–61). We postu-
late that the decrease of total thiols and glutathione levels in
mutant cells may be a consequence of the consumption of glu-
tathione by a process such as protein glutathionylation.
HPLC experiments (Table 1) show NADPH levels in
mutant cells being one quarter that in the wild-type, associated
with a shift of the NADPH/NADPþratio from 1.9 in wild-type
cells to 0.6 in frataxin-deficient cells. Indeed, due to the
significantly higher Gpx activity (Fig. 2), GSSG is presumably
reduced back to GSH by glutathione reductase, using NADPH
for a reducing power, so as to maintain the redox status of the
cells. NADPH provides electrons for the reduction of both
the oxidized disulphide form of thioredoxin, catalysed by
thioredoxin reductase, and GSSG to GSH catalysed by gluta-
thione reductase (8,62). The substantial imbalanced NADPH/
NADPþratio we observed confirms the severe oxidative
stress status and the malfunction of the thiol-dependent anti-
oxidant defences of mutant cells, suggesting alternative
metabolic mechanisms are used to survive the stress associated
with frataxin deficiency. Surprisingly, our results show a
decrease of the total (NADPHþNADPþ) levels, and this
implies oxidative modification of NADPH, such as the
Human Molecular Genetics, 2008, Vol. 17, No. 18 2797
by guest on June 5, 2013
degradation into NADH catalysed by phosphatases. Supporting
this hypothesis, the genes coding for glycerol-3-phosphatases,
RHR2 and HOR2, are induced in frataxin-deficient cells
(microarray data, not shown).
In S. cerevisiae cells, glucose-6-phosphate dehydrogenase
and, to a lesser extent, NADPþ-specific IDP are believed to
be the major sources of the NADPH used by thiol-dependent
peroxidases in the cytosol (46). In frataxin-deficient cells,
specific G6PDH activity was significantly higher than wild-
type (three times) and the expression of ZWF1, coding for
G6PDH, was induced (Table 2). In addition, microarray data
(not shown) indicate that the expression of IDP1,2,3 coding
for IDPs is two or three times higher than in the wild-type. Con-
sequently, the pentose phosphate pathway is stimulated in
mutant cells, as by the NADPH/NADPþratios. Indeed, when
a cell is rapidly converting NADPH to NADPþ, as observed
in frataxin-deficient cells, the level of NADPþrises, allos-
terically stimulating G6PDH to accelerate NADPH biosyn-
thesis. However, the strong induction of G6PDH cannot
restore the NADPH/NADPþpools sufficiently to be able to
maintain the normal redox status of the cells. Our results are
in good agreement with previous work showing that G6PDH
expression is enhanced by oxidative stress (63). Moreover, it
has been reported that this activation of the enzyme is associ-
ated with a decrease of the glutathione pool and that this
increase is blocked by treatment with antioxidants that specifi-
cally replenish the intracellular GSH (49,50). G6PDH
expression is inversely correlated to intracellular GSH levels
and GSH replenishment may depend on G6PDH expression
(49); this is consistent with the induction of ZWF1 we detected
by the transcriptional analysis. Our results confirm the unusual
metabolism, which could be relevant to defects in glucose
metabolism in FA patients (64,65).
Iron accumulation inside the mitochondria is one of the
main phenotypes observed in frataxin-deficient cells and has
been described as being responsible for the severe oxidative
stress status of these cells due to Fenton chemistry processes
(reviewed in 3). In addition, GSH depletion causes a specific
defect in the maturation of iron–sulphur proteins and cellular
iron accumulation (17). We studied the glutathione-dependent
response of wild-type and frataxin-deficient cells to iron-
dependent stress conditions.
When cultured in the presence of 100-times excess ferric
citrate, total glutathione levels and both specific Gpx and
G6PDH activities returned to values similar to those in wild-
type cells (Fig. 5). It, therefore, appears that, in the presence
of an excess iron, frataxin-deficient cells are able to restore
the initial thiol-dependent antioxidant status. However, there
were little changes at the transcriptional levels (Table 2).
The increase of total glutathione levels cannot be attributed
to an increased synthesis, because at least GSH1 and GSH2
transcription was not affected by iron addition (Table 2).
In yeast, high-affinity iron uptake is regulated at the tran-
scriptional level by Aft1. Aft1-dependent activation of the
‘iron regulon’ is constitutive in Dyfh1 mutants such that the
high affinity iron transport system of the cell membrane is
strongly induced regardless of the iron content of the
medium (52). In Dyfh1 mutants, the cytosolic iron concen-
tration is lower than in wild-type cells grown in the same
medium, but it accumulates in the mitochondria. Our results
suggest that an excess iron in the culture may allow to
restore a normal glutathione-dependent redox status, influen-
We report the first dynamic study of the specific
glutathione-dependent adaptative response to iron-dependent
stress conditions. Wild-type and frataxin-deficient cells
reacted differently to the stress induced by the addition of
iron in the medium (Figs. 6 and 7). For wild-type cells, trans-
fer to an iron-supplemented culture medium was followed by a
significant decrease of glutathione levels within 1 min, fol-
lowed by adaptation and equilibration within 5 min and a
return to initial conditions. In parallel, specific Gpx activity
increased then returned to the initial values after 10 min.
Thus, there was a decrease in total glutathione associated
with an increase in Gpx activity, which consumes GSH to
form GSSG. In contrast, exposure of mutant cells to excess
iron did not affect the glutathione levels or the Gpx and
G6PDH activities; excess iron does not appear to act as an
additional stress for mutant cells which already accumulate
iron inside the mitochondria.
Wild-type cells were also stressed by transfer into iron-
depleted conditions. The transfer of the cells to a Bio101
culture medium was followed by an increase of total gluta-
thione levels, without a subsequent return to initial conditions;
the cells are thus unable to continue their adaptative response
in condition of severe iron deficiency. Once again, the
glutathione-dependent redox status of mutant cells was not
affected by iron deficiency. Indeed, mutant cells, whereas
accumulating iron, might behave like iron-deficient cells.
Our findings show that wild-type cells show an acute response
to iron-stress conditions due to a change in the redox potential
of the cellular environment, whereas mutant cells, which are
constitutively stressed and are able to accumulate iron inside
the mitochondria, had already responded to long-term
iron-excess conditions. As suggested by studies in gsh1
mutants (17) and more recently by Wheeler and Grant (10),
it appears likely that the regulation of GSH and iron meta-
bolism is intrinsically linked. Indeed, GSH and glutaredoxin 3
and 4 are involved in iron sensing by the Aft1 transcriptional
regulator of iron homeostasis (66,67).
In conclusion, we report evidence of an impairment of glu-
tathione homeostasis in a yeast model of Friedreich’s ataxia,
revealing this molecule as an indicator of the redox status of
frataxin-deficient cells, in agreement with the role of free rad-
icals in the pathophysiology of the disease. Our findings indi-
cate that glutathione assays could be use as a routine tool to
detect the oxidative stress status of frataxin-deficient cells.
We also describe a strong link between the glutathione-
dependent redox status of the cells and iron pathways, provid-
ing new insights into the complicated and controversial
relationship between iron and oxidative stress in the study of
MATERIALS AND METHODS
Yeast strains, media and growth conditions
The S. cerevisiae strains used were S150-2B (MATa, his3-D1,
leu2-3112, trp1-289, ura3-52), S150-2BDyfh1 (Dyfh1::TRP1),
2798 Human Molecular Genetics, 2008, Vol. 17, No. 18
by guest on June 5, 2013
(MATa, ura3-52, lys2-801, ade2-101, trp1D63, his3D200,
leu2D1, cyh2) and the yfh1 shuffle strain YPH499 [Dyfh1::TRP1
(pRS318-LEU2-CYH2-YFH1)] (68). In this strain, the yfh1
mutation is covered by pRS318, a plasmid containing the CEN
and CYH2 and the YFH1 HindIII genomic fragment. The
plasmid was removed prior to experiments by counter selection
on medium containing 10 mg/ml cycloheximide, which is toxic
in the presence of the CYH2 allele, under anaerobic conditions.
The resulting mutant was named YPH499Dyfh1.
Unless otherwise stated, cells were grown at 308C in
minimal YNB (yeast nitrogen base, Bio 101, Inc., 2%
D-glucose) plus the required amino acids. Other media used
were complete YPD (1% yeast extract, 2% bacteriological
peptone and 2% D-glucose) and YPR (1% yeast extract, 1%
bacteriological peptone, 2% raffinose and 0.2% glucose).
Minimal YNB (yeast nitrogen base without copper and iron,
Bio 101, Inc., 2% glucose) medium supplemented with the
required amino acids and 4 mM copper sulphate was used
for cultures in iron-deficient medium. Iron-rich medium was
supplemented with 100 mM iron citrate. For growth medium
shift experiments, yeast strains were cultured to early log
phase and the cells collected by filtration. Cells were collected
from the culture media at various times after resuspension in
the new medium.
Preparation of crude cell homogenates and measurement
of protein content
Yeast cells were cultured until reaching an OD600nm?0.7 and
harvested by centrifugation. The pellets were resuspended in
50 mM potassium phosphate buffer pH 7.8 in the presence of
protease inhibitors and the cells were disrupted using glass
beads, and centrifuged for 30 min at 5000 g; and the super-
natant was used as the crude cell extract.
Protein content of crude cell homogenates or isolated mito-
chondria was determined using the BCA (bicinchoninic acid)
assay (69) and enzyme activities and thiol contents are
reported in units/mg protein.
Isolation of yeast mitochondria
Mitochondria were isolated as described by Gasser and Schatz
(70) with modifications. Yeast cells were cultured in 1.5 l of
YPR medium and harvested by centrifugation. They were
then incubated with 2 mg of Zymolyase-100T (MD) per
gram of cells in spheroblast buffer (1.2 M sorbitol, 50 mM
Tris pH 7.5, 10 mM DTT) at 308C for 60 min. The spheroblasts
were washed twice with the spheroblast buffer without DTT,
suspended in ice-cold homogenization buffer (20 mM Hepes-
KOH pH 7.4, 0.6 M sorbitol) to a concentration of 0.8 g of
cells/ml and then homogenized in a Dounce homogenizer
(Kontes Glass Co., Vineland, NJ, USA). The homogenate
(45 ml) was diluted with an equal volume of the homogeni-
zation buffer and centrifuged at 1500 g for 5 min at 48C.
The supernatant was saved and the pellet was homogenized
in 40 ml of homogenization buffer, and re-centrifuged as
above. The two supernatants were combined and centrifuged
again as before to remove residual cell debris. Mitochondria
were collected from the supernatant by centrifugation
(10 min, 9600 g, 48C) and resuspended in a small volume
(1–2 ml) of homogenization buffer.
Measurement of total reduced thiol content
Thiol content was determined by spectrophotometric quantifi-
cation of the conversion of 5.50-dithiobis-2-nitrobenzoı ¨c acid
(DTNB) into 5-thio-2-nitrobenzoı ¨c acid (TNB) at 412 nm
(71). Standard curves were obtained using various GSH and/
or cystein concentrations. All data points in the figures and
the values listed, expressed as nmoles of total reduced
thiols/mg of protein, are means of at least three independent
Determination of glutathione levels (GSH1GSSG)
Glutathione levels were determined using a modified pro-
cedure of the recycling enzymatic assay described by Tietze
(72). Yeast strains were grown in minimum medium and
cells were harvested at the appropriate growth phase by cen-
trifugation. Extracellular glutathione was measured directly
in the resulting supernatant. For the estimation of total intra-
cellular glutathione, cell pellets were washed and resuspended
in 50 mM potassium phosphate buffer pH 7.8 containing
ice-cold 5% 5-sulphosalicylic acid, and broken with glass
beads. The resulting mixture was clarified by centrifugation
(30 min, 5000 g, 48C) and the supernatant was used to deter-
mine total free glutathione. A typical reaction mixture con-
tained the cell extract, 20 mM DTNB and 10 mM NADPH in
50 mM potassium phosphate buffer pH 7.8. The reaction was
started by addition of glutathione reductase (1.5 units/ml)
and the kinetics of conversion of DTNB into TNB were fol-
lowed spectrophotometrically at 412 nm. Glutathione concen-
trations were calculated from standard curves obtained with
various GSH and GSSG concentrations, using the rates of
TNB formation, and are expressed as nmoles of glutathione/
mg of protein. All data points in the figures and the values
listed are means of at least three determinations, and Student’s
t-test was used to identify significant differences. For quantifi-
cation of oxidized glutathione (GSSG), samples (including
2-vinylpyridine for 1 h at room temperature before analysis.
Measurement of glutathione peroxidase activity
Glutathione peroxidase (Gpx) activity in cell extracts and iso-
lated mitochondria was assayed as previously described
(40,41,43) with modifications, using tert-butyl hydroperoxide
(t-BHP) as a substrate for the peroxidase. The assay was per-
formed in 50 mM potassium phosphate buffer pH 7.0 contain-
ing 10 mM reduced GSH, 200 mM NADPH and 4 mM t-BHP.
The reaction was started by addition of glutathione reductase
to a final concentration of 0.45 units/ml, and the kinetics of
NADPH oxidation were followed spectrophotometrically at
340 nm. Specific glutathione peroxidase activity was then
e340nm¼ 6200 M21cm21. One unit of glutathione peroxidase
activity was defined as the amount of enzyme which catalyses
oxidation of 1 mmole of GSH into GSSG per minute. All data
points in the figures and the values listed are means of three
Human Molecular Genetics, 2008, Vol. 17, No. 18 2799
by guest on June 5, 2013
or more determinations and all results are reported relative to
the protein concentration of the cell extract.
Measurement of glucose-6-phosphate
Glucose-6-phosphate dehydrogenase (G6PD) activity was deter-
mined as described in references (73,74). G6PDH catalyses the
conversion of glucose-6-phosphate to 6-phosphate-gluconolac-
enzyme of the pentose phosphate pathway. Because 6PGD also
(G6PDþ6PGD) were measured separately as described by Tian
et al. (74), to determine enzyme activity accurately. The conver-
sion of NADPþto NADPH catalysed by the two dehydrogenase
enzymes was measured spectrophotometrically by following the
increase of absorbance at 340 nm due to the conversion of
NADPþto NADPH. 6PGD enzymatic activity was measured
firstin a reaction mixturecontaining 50 mM potassium phosphate
buffer pH 7.8 (containing 1 mM MgCl), 0.2 mM NADPþand
0.4 mM 6-phosphogluconate. The kinetics of NADPH production
by the 6PGD reaction were measured, and glucose-6-phosphate
(the specific substrate for G6PD) was then added to the cuvette
to 1 mM final concentration, resulting in an increase of the
rate of production of NADPH, corresponding to the total
dehydrogenase activity. G6PD activity was then calculated by
subtracting the activity of 6PGD from total dehydrogenase
activity and the results are expressed as nmoles of NADPH/mg
Acid extraction was used for oxidized nicotinamide nucleo-
tides (NADPþ) and alkaline extraction for reduced forms
(NADPH), using a procedure modified from references
(75,76). After protein extraction with 50% TCA and centrifu-
gation for 5 min (6000 g), the resulting supernatants were
treated either with 0.5 M HClO4(35% v/v in ethanol) for
acid extraction, or with 0.5 M KOH (50% in ethanol) for
alkaline extraction. The samples were homogenized, incubated
for 10 min on ice and centrifuged for 15 min at 48C (5000 g).
For maximal recovery of reduced pyridine nucleotides,
alkaline extracts were incubated for a further 10 min at
608C, and then allowed to cool for 10 min in ice, as previously
described in Klein et al. (75). Before assaying the nucleotide
extracts, the supernatants were neutralized to pH 7.0 by
carefully adding 10 N HCl to the alkaline extracts, and
1 M KOH to acid extracts, and thoroughly vortexing. The
neutralized extracts were then kept at 08C for at least 10 min
before centrifugation for 5 min at 5000 g. The resulting
samples were then injected on the same day into the HPLC
HPLC analysis of NADPH/NADP1pools
Phase-HPLC (Dionex HPLC system interfaced with the
Kromasil ODS2 C18 column (length ¼ 250 nm, internal
nucleotideswere analysed by Reverse
diameter ¼ 4 mm, particle size ¼ 5 mm) (Interchim) at room
temperature. The mobile phase used for the separation of
these nucleotides consisted of two eluants: solvent A was
10 mM ammonium acetate buffer (pH 6.0) and solvent B was
methanol. Compounds were separated by the following dis-
continuous gradient at a flow rate of 0.8 ml/min: a linear
increase in solvent B to 25% over 20 min, followed by an
increase to 30% in the next 2 min, and stable at 30% for
4 min; this was followed by a decrease from 30 to 0% over
the next minute, and the initial conditions were then main-
tained for 20 min. The products were monitored spectrophoto-
metrically at 260 and 340 nm and quantified by integration of
the peak absorbance area, employing a calibration curve estab-
lished with various known concentrations of NADPH and
Iron accumulation measurements
Iron accumulation by the cells was measured in microtitre
plates, after growing the cells overnight in minimum
medium with 1 to 100 mM
described previously (77).
55Fe(II)-citrate (86 mCi/mg), as
RNA isolation and real-time quantitative PCR analysis
Total RNA was extracted from cells cultured in YNB media to
an OD600nm?0.7 using the hot phenol method as described pre-
ing to the kit manufacturer’s instructions (First Strand cDNA
Synthesis Kit for RT–PCR, Roche Diagnostics). PCRs were
performed on a LightCyclerw480 System (Roche Diagnostics)
in 384-well plates. Each reaction was carried out in 10 ml with
to5 ngofreversetranscribedRNA),250or350 nMprimercon-
centration depending on the primer pair and 0.8X ABsoluteTM
QPCR SYBRwGreen Mix (ABgene Inc.). Serial dilutions of
ive PCR standard curve. The LightCycler protocol was: 15 min
tion for 15 s, 608C annealing for 25 s and 728C elongation for
10 s; and melting at 958C for 5 s, 708C for 60 s, and then
heating to 958C. Water was used as the template for negative
control amplifications included with each PCR run. All reac-
tions were performed in duplicate in each 384 well plate. Data
were analysed using the Roche LightCyclerw480 software
and CP was calculated by the Second Derivate Maximum
Method. The amount of the target mRNA was examined and
normalized to the RPO21 gene mRNA. The relative expression
ratio of a target gene was calculated as described by Pfaffl (79),
based on real-time PCR efficiencies. Primer pairs are listed in
Table 2 and were designed with Primer3 software to generate
products of 82 to 150 bp. Results reported were obtained from
at least three biological replicates and PCR runs were repeated
at least twice.
500 ng ofRNA, using
Conflict of Interest statement. None declared.
2800Human Molecular Genetics, 2008, Vol. 17, No. 18
by guest on June 5, 2013
This work has been funded by the ‘Agence Nationale de La
Recherche: Maladies Rares (french government)’. ANR-06-
1. Campuzano, V., Montermini, L., Lutz, Y., Cova, L., Hindelang, C.,
Jiralerspong, S., Trottier, Y., Kish, S.J., Faucheux, B., Trouillas, P. et al.
(1997) Frataxin is reduced in Friedreich ataxia patients and is associated
with mitochondrial membranes. Hum. Mol. Genet., 6, 1771–1780.
2. Campuzano, V., Montermini, L., Molto, M.D., Pianese, L., Cossee, M.,
Cavalcanti, F., Monros, E., Rodius, F., Duclos, F., Monticelli, A. et al.
(1996) Friedreich’s ataxia: autosomal recessive disease caused by an
intronic GAA triplet repeat expansion. Science, 271, 1423–1427.
3. Pandolfo, M. (2006) Iron and Friedreich ataxia. J. Neural. Transm. Suppl.,
4. Napoli, E., Taroni, F. and Cortopassi, G.A. (2006) Frataxin, iron–sulfur
clusters, heme, ROS, and aging. Antioxid. Redox Signal, 8, 506–516.
5. Piemonte, F., Pastore, A., Tozzi, G., Tagliacozzi, D., Santorelli, F.M.,
Carrozzo, R., Casali, C., Damiano, M., Federici, G. and Bertini, E. (2001)
Glutathione in blood of patients with Friedreich’s ataxia. Eur. J. Clin.
Invest., 31, 1007–1011.
6. Tozzi, G., Nuccetelli, M., Lo Bello, M., Bernardini, S., Bellincampi, L.,
Ballerini, S., Gaeta, L.M., Casali, C., Pastore, A., Federici, G. et al. (2002)
Antioxidant enzymes in blood of patients with Friedreich’s ataxia. Arch.
Dis. Child., 86, 376–379.
7. Herrero, E., Ros, J., Belli, G. and Cabiscol, E. (2007) Redox control and
oxidative stress in yeast cells. Biochim. Biophys. Acta, 64, 1518–1530.
8. Holmgren, A., Johansson, C., Berndt, C., Lonn, M.E., Hudemann, C. and
Lillig, C.H. (2005) Thiol redox control via thioredoxin and glutaredoxin
systems. Biochem. Soc. Trans., 33, 1375–1377.
9. Toledano, M.B., Kumar, C., Le Moan, N., Spector, D. and Tacnet, F.
(2007) The system biology of thiol redox system in Escherichia coli and
yeast: differential functions in oxidative stress, iron metabolism and DNA
synthesis. FEBS Lett., 581, 3598–3607.
10. Wheeler, G.L. and Grant, C.M. (2004) Regulation of redox homeostasis in
the yeast Saccharomyces cerevisiae. Physiol. Plant., 120, 12–20.
11. Penninckx, M.J. (2002) An overview on glutathione in Saccharomyces
versus non-conventional yeasts. FEMS Yeast Res., 2, 295–305.
12. Perrone, G.G., Grant, C.M. and Dawes, I.W. (2005) Genetic and
environmental factors influencing glutathione homeostasis in
Saccharomyces cerevisiae. Mol. Biol. Cell, 16, 218–230.
13. Pocsi, I., Prade, R.A. and Penninckx, M.J. (2004) Glutathione, altruistic
metabolite in fungi. Adv. Microb. Physiol., 49, 1–76.
14. Hammond, C.L., Lee, T.K. and Ballatori, N. (2001) Novel roles for
glutathione in gene expression, cell death, and membrane transport of
organic solutes. J. Hepatol., 34, 946–954.
15. Vuilleumier, S., Sorribas, H. and Leisinger, T. (1997) Identification of a
novel determinant of glutathione affinity in dichloromethane
dehalogenases/glutathione S-transferases. Biochem. Biophys. Res.
Commun., 238, 452–456.
16. Rodriguez-Manzaneque, M.T., Tamarit, J., Belli, G., Ros, J. and
Herrero, E. (2002) Grx5 is a mitochondrial glutaredoxin required for the
activity of iron/sulfur enzymes. Mol. Biol. Cell, 13, 1109–1121.
17. Sipos, K., Lange, H., Fekete, Z., Ullmann, P., Lill, R. and Kispal, G.
(2002) Maturation of cytosolic iron–sulfur proteins requires glutathione.
J. Biol. Chem., 277, 26944–26949.
18. Herrero, E. and de la Torre-Ruiz, M.A. (2007) Monothiol glutaredoxins: a
common domain for multiple functions. Cell Mol. Life Sci., 64, 1518–1530.
19. Jamieson, D.J. (1998) Oxidative stress responses of the yeast
Saccharomyces cerevisiae. Yeast, 14, 1511–1527.
20. Oakley, A.J. (2005) Glutathione transferases: new functions. Curr. Opin.
Struct. Biol., 15, 716–723.
21. Miranda-Vizuete, A., Damdimopoulos, A.E. and Spyrou, G. (2000) The
mitochondrial thioredoxin system. Antioxid. Redox Signal, 2, 801–810.
22. Pedrajas, J.R., Kosmidou, E., Miranda-Vizuete, A., Gustafsson, J.A.,
Wright, A.P. and Spyrou, G. (1999) Identification and functional
characterization of a novel mitochondrial thioredoxin system in
Saccharomyces cerevisiae. J. Biol. Chem., 274, 6366–6373.
23. Trotter, E.W. and Grant, C.M. (2002) Thioredoxins are required for
protection against a reductive stress in the yeast Saccharomyces
cerevisiae. Mol. Microbiol., 46, 869–878.
24. Jeong, J.S., Kwon, S.J., Kang, S.W., Rhee, S.G. and Kim, K. (1999)
Purification and characterization of a second type thioredoxin peroxidase
(type II TPx) from Saccharomyces cerevisiae. Biochemistry, 38, 776–783.
25. Park, S.G., Cha, M.K., Jeong, W. and Kim, I.H. (2000) Distinct
physiological functions of thiol peroxidase isoenzymes in Saccharomyces
cerevisiae. J. Biol. Chem., 275, 5723–5732.
26. Lee, J.C., Straffon, M.J., Jang, T.Y., Higgins, V.J., Grant, C.M. and
Dawes, I.W. (2001) The essential and ancillary role of glutathione in
Saccharomyces cerevisiae analysed using a grande gsh1 disruptant strain.
FEMS Yeast Res., 1, 57–65.
27. Muller, E.G. (1996) A glutathione reductase mutant of yeast accumulates
high levels of oxidized glutathione and requires thioredoxin for growth.
Mol. Biol. Cell, 7, 1805–1813.
28. Grant, C.M., MacIver, F.H. and Dawes, I.W. (1996) Glutathione is an
essential metabolite required for resistance to oxidative stress in the yeast
Saccharomyces cerevisiae. Curr. Genet., 29, 511–515.
29. Grant, C.M., Perrone, G. and Dawes, I.W. (1998) Glutathione and catalase
provide overlapping defenses for protection against hydrogen peroxide in
the yeast Saccharomyces cerevisiae. Biochem. Biophys. Res. Commun.,
30. Izawa, S., Inoue, Y. and Kimura, A. (1995) Oxidative stress response in
yeast: effect of glutathione on adaptation to hydrogen peroxide stress in
Saccharomyces cerevisiae. FEBS Lett., 368, 73–76.
31. Madeo, F., Frohlich, E., Ligr, M., Grey, M., Sigrist, S.J., Wolf, D.H. and
Frohlich, K.U. (1999) Oxygen stress: a regulator of apoptosis in yeast.
J. Cell. Biol., 145, 757–767.
32. Costa, V. and Moradas-Ferreira, P. (2001) Oxidative stress and signal
transduction in Saccharomyces cerevisiae: insights into ageing, apoptosis
and diseases. Mol. Aspects Med., 22, 217–246.
33. Moradas-Ferreira, P. and Costa, V. (2000) Adaptive response of the yeast
Saccharomyces cerevisiae to reactive oxygen species: defences, damage
and death. Redox Rep., 5, 277–285.
34. Nestelbacher, R., Laun, P., Vondrakova, D., Pichova, A., Schuller, C. and
Breitenbach, M. (2000) The influence of oxygen toxicity on yeast mother
cell-specific aging. Exp. Gerontol., 35, 63–70.
35. Schultz, D. and Harrison, D.G. (2000) Quest for fire: seeking the source of
pathogenic oxygen radicals in atherosclerosis. Arterioscler. Thromb. Vasc.
Biol., 20, 1412–1413.
36. Grant, C.M. (2001) Role of the glutathione/glutaredoxin and thioredoxin
systems in yeast growth and response to stress conditions. Mol.
Microbiol., 39, 533–541.
37. Stephen, D.W. and Jamieson, D.J. (1996) Glutathione is an important
antioxidant molecule in the yeast Saccharomyces cerevisiae. FEMS
Microbiol. Lett., 141, 207–212.
38. Bursell, S.E. and King, G.L. (2000) The potential use of glutathionyl
hemoglobin as a clinical marker of oxidative stress. Clin. Chem., 46,
39. Calabrese, V., Lodi, R., Tonon, C., D’Agata, V., Sapienza, M., Scapagnini,
G., Mangiameli, A., Pennisi, G., Stella, A.M. and Butterfield, D.A. (2005)
Oxidative stress, mitochondrial dysfunction and cellular stress response
in Friedreich’s ataxia. J. Neurol. Sci., 233, 145–162.
40. Galiazzo, F., Schiesser, A. and Rotilio, G. (1987) Glutathione peroxidase
in yeast. Presence of the enzyme and induction by oxidative conditions.
Biochem. Biophys. Res. Commun., 147, 1200–1205.
41. Avery, A.M. and Avery, S.V. (2001) Saccharomyces cerevisiae expresses
three phospholipid hydroperoxide glutathione peroxidases. J. Biol. Chem.,
42. Avery, A.M., Willetts, S.A. and Avery, S.V. (2004) Genetic dissection of
the phospholipid hydroperoxidase activity of yeast Gpx3 reveals its
functional importance. J. Biol. Chem., 279, 46652–46658.
43. Inoue, Y., Matsuda, T., Sugiyama, K., Izawa, S. and Kimura, A. (1999)
Genetic analysis of glutathione peroxidase in oxidative stress response of
Saccharomyces cerevisiae. J. Biol. Chem., 274, 27002–27009.
44. Kho, C.W., Lee, P.Y., Bae, K.H., Cho, S., Lee, Z.W., Park, B.C., Kang, S.,
Lee do, H. and Park, S.G. (2006) Glutathione peroxidase 3 of
Saccharomyces cerevisiae regulates the activity of methionine sulfoxide
reductase in a redox state-dependent way. Biochem. Biophys. Res.
Commun., 348, 25–35.
45. Lee, P.Y., Kho, C.W., Lee do, H., Kang, S., Kang, S., Lee, S.C., Park, B.C.,
Cho, S., Bae, K.H. and Park, S.G. (2007) Glutathione peroxidase 3 of
Human Molecular Genetics, 2008, Vol. 17, No. 182801
by guest on June 5, 2013
Saccharomyces cerevisiae suppresses non-enzymatic proteolysis of
glutamine synthetase in an activity-independent manner. Biochem. Biophys.
Res. Commun., 362, 405–409.
46. Minard, K.I. and McAlister-Henn, L. (1999) Dependence of peroxisomal
beta-oxidation on cytosolic sources of NADPH. J. Biol. Chem., 274,
47. Minard, K.I. and McAlister-Henn, L. (2001) Antioxidant function of
cytosolic sources of NADPH in yeast. Free Radic. Biol. Med., 31, 832–843.
48. Minard, K.I., Carroll, C.A., Weintraub, S.T. and Mc-Alister-Henn, L.
(2007) Changes in disulfide bond content of proteins in a yeast strain
lacking major sources of NADPH. Free Radic. Biol. Med., 42, 106–117.
49. Salvemini, F., Franze, A., Iervolino, A., Filosa, S., Salzano, S. and
Ursini, M.V. (1999) Enhanced glutathione levels and oxidoresistance
mediated by increased glucose-6-phosphate dehydrogenase expression.
J. Biol. Chem., 274, 2750–2757.
50. Ursini, M.V., Parrella, A., Rosa, G., Salzano, S. and Martini, G. (1997)
Enhanced expression of glucose-6-phosphate dehydrogenase in human
cells sustaining oxidative stress. Biochem. J., 323, 801–806.
51. Srikanth, C.V., Vats, P., Bourbouloux, A., Delrot, S. and Bachhawat, A.K.
(2005) Multiple cis-regulatory elements and the yeast sulphur regulatory
network are required for the regulation of the yeast glutathione
transporter, Hgt1p. Curr. Genet., 47, 345–358.
52. Babcock, M., de Silva, D., Oaks, R., Davis-Kaplan, S., Jiralerspong, S.,
Montermini, L., Pandolfo, M. and Kaplan, J. (1997) Regulation of
mitochondrial iron accumulation by Yfh1p, a putative homolog of
frataxin. Science, 276, 1709–1712.
53. Foury, F. and Cazzalini, O. (1997) Deletion of the yeast homologue of the
human gene associated with Friedreich’s ataxia elicits iron accumulation
in mitochondria. FEBS Lett., 411, 373–377.
54. Adamis, P.D., Gomes, D.S., Pinto, M.L., Panek, A.D. and Eleutherio, E.C.
(2004) The role of glutathione transferases in cadmium stress. Toxicol.
Lett., 154, 81–88.
55. Castro,F.A.,Herdeiro,R.S., Panek,A.D.,Eleutherio,E.C.andPereira,M.D.
(2007) Menadione stress inSaccharomyces cerevisiaestrainsdeficientinthe
glutathione transferases. Biochim. Biophys. Acta, 1770, 213–220.
56. Collinson, E.J., Wheeler, G.L., Garrido, E.O., Avery, A.M., Avery, S.V.
and Grant, C.M. (2002) The yeast glutaredoxins are active as glutathione
peroxidases. J. Biol. Chem., 277, 16712–16717.
57. Bourbouloux, A., Shahi, P., Chakladar, A., Delrot, S. and Bachhawat, A.K.
(2000) Hgt1p, a high affinity glutathione transporter from the yeast
Saccharomyces cerevisiae. J. Biol. Chem., 275, 13259–13265.
58. Coan, C., Ji, J.Y., Hideg, K. and Mehlhorn, R.J. (1992) Protein sulfhydryls
are protected from irreversible oxidation by conversion to mixed
disulfides. Arch. Biochem. Biophys., 295, 369–378.
59. Cotgreave, I.A., Weis, M., Berggren, M., Sandy, M.S. and Moldeus, P.W.
(1988) Determination of the intracellular protein thiol distribution of
hepatocytes using monobromobimane derivatisation of intact cells and
isolated subcellular fractions. J. Biochem. Biophys. Methods, 16, 247–254.
60. Shenton, D. and Grant, C.M. (2003) Protein S-thiolation targets glycolysis
and protein synthesis in response to oxidative stress in the yeast
Saccharomyces cerevisiae. Biochem. J., 374, 513–519.
61. Thomas, J.A., Poland, B. and Honzatko, R. (1995) Protein sulfhydryls and
their role in the antioxidant function of protein S-thiolation. Arch.
Biochem. Biophys., 319, 1–9.
62. Herrero, E., Ros, J., Tamarit, J. and Belli, G. (2006) Glutaredoxins in
fungi. Photosynth. Res., 89, 127–140.
63. Inoue, Y., Sugiyama, K., Izawa, S. and Kimura, A. (1998) Molecular
identification of glutathione synthetase (GSH2) gene from Saccharomyces
cerevisiae. Biochim. Biophys. Acta, 1395, 315–320.
64. Finocchiaro, G., Baio, G., Micossi, P., Pozza, G. and di Donato, S. (1988)
Glucose metabolism alterations in Friedreich’s ataxia. Neurology, 38,
65. Ristow, M. (2004) Neurodegenerative disorders associated with diabetes
mellitus. J. Mol. Med., 8, 510–529.
66. Ojeda, L., Keller, G., Muhlenhoff, U., Rutherford, J.C., Lill, R. and
Winge, D.R. (2006) Role of glutaredoxin-3 and glutaredoxin-4 in the iron
regulation of the Aft1 transcriptional activator in Saccharomyces
cerevisiae. J. Biol. Chem., 281, 17661–17669.
(2006) Glutaredoxins Grx3 and Grx4 regulate nuclear localisation of
Aft1 and the oxidative stress response in Saccharomyces cerevisiae.
J. Cell Sci., 119, 4554–4564.
68. Zhang, Y., Lyver, E.R., Knight, S.A., Lesuisse, E. and Dancis, A. (2005)
Frataxin and mitochondrial carrier proteins, Mrs3p and Mrs4p, cooperate
in providing iron for heme synthesis. J. Biol. Chem., 280, 19794–19807.
69. Smith, P.K., Krohn, R.I., Hermanson, G.T., Mallia, A.K., Gartner, F.H.,
(1985) Measurement of protein using bicinchoninic acid. Anal. Biochem.,
70. Gasser, S.M. and Schatz, G. (1983) Import of proteins into mitochondria.
In vitro studies on the biogenesis of the outer membrane. J. Biol. Chem.,
71. Sedlak, J. and Lindsay, R.H. (1968) Estimation of total, protein-bound,
and nonprotein sulfhydryl groups in tissue with Ellman’s reagent. Anal.
Biochem., 25, 192–205.
72. Tietze, F. (1969) Enzymic method for quantitative determination of
nanogram amounts of total and oxidized glutathione: applications to
mammalian blood and other tissues. Anal. Biochem., 27, 502–522.
73. Battistuzzi, G., D’Urso, M., Toniolo, D., Persico, G.M. and Luzzatto, L.
(1985) Tissue-specific levels of human glucose-6-phosphate
dehydrogenase correlate with methylation of specific sites at the 3’ end of
the gene. Proc. Natl Acad. Sci. USA, 82, 1465–1469.
74. Tian, W.N., Pignatare, J.N. and Stanton, R.C. (1994) Signal transduction
proteins that associate with the platelet-derived growth factor (PDGF)
receptor mediate the PDGF-induced release of glucose-6-phosphate
dehydrogenase from permeabilized cells. J. Biol. Chem., 269, 14798–
75. Klein, A., Chan, A.W., Caplan, B.U. and Malin, A. (1990) NADPþ
reduction by human lymphocytes. Clin. Exp. Immunol., 82, 170–173.
76. Mailinger, W., Baumeister, A., Reuss, M. and Rizzi, M. (1998) Rapid and
highly automated determination of adenine and pyridine nucleotides in
extracts of Saccharomyces cerevisiae using a micro robotic sample
preparation-HPLC system. J. Biotechnol., 63, 155–166.
77. Bulteau, A.L., Dancis, A., Gareil, M., Montagne, J.J., Camadro, J.M. and
Lesuisse, E. (2007) Oxidative stress and protease dysfunction in the yeast
model of Friedreich ataxia. Free Radic. Biol. Med., 42, 1561–1570.
78. Kohrer, K. and Domdey, H. (1991) Preparation of high molecular weight
RNA. Methods Enzymol., 194, 398–405.
79. Pfaffl, M.W. (2001) A new mathematical model for relative quantification
in real-time RT–PCR. Nucleic Acids Res., 29, 45–50.
2802 Human Molecular Genetics, 2008, Vol. 17, No. 18
by guest on June 5, 2013